Skip to main page content

• HOME
• Current Issue
• Archive
• Alerts
• About ASM
• Contact us
• Tech Support
• Journals.ASM.org

This Article Abstract Full Text (PDF) Alert me when this article is cited Alert me if a correction is posted Services Similar articles in this journal Similar articles in PubMed Alert me to new issues of the journal Download to citation manager Reprints and Permissions Copyright Information Books from ASM Press MicrobeWorld Citing Articles Citing Articles via HighWire Citing Articles via Google Scholar Google Scholar Articles by Rasmussen, J. W. Articles by Benach, J. L. Search for Related Content PubMed PubMed Citation Articles by Rasmussen, J. W. Articles by Benach, J. L.

 Previous Article  |  Next Article 

Infection and Immunity, December 2006, p. 6590-6598, Vol. 74, No. 12
0019-9567/06/$08.00+0     doi:10.1128/IAI.00868-06
Copyright © 2006, American Society for Microbiology. All Rights Reserved.

Mac-1⁺ Cells Are the Predominant Subset in the Early Hepatic Lesions of Mice Infected with Francisella tularensis

John W. Rasmussen,¹ Jeronimo Cello,¹ Horacio Gil,^1, Colin A. Forestal,¹ Martha B. Furie,^1,2,3 David G. Thanassi,^1,3 and Jorge L. Benach^1,3*

Center for Infectious Diseases,¹ Department of Pathology,² Department of Molecular Genetics and Microbiology, Stony Brook University, Stony Brook, New York 11794³

Received 31 May 2006/ Returned for modification 7 July 2006/ Accepted 17 September 2006

Top Abstract Introduction Materials and Methods Results and Discussion References

 
The cell composition of early hepatic lesions of experimental murine tularemia has not been characterized with specific markers. The appearance of multiple granulomatous-necrotic lesions in the liver correlates with a marked increase in the levels of serum alanine transferase and lactate dehydrogenase. Francisella tularensis, detected by specific antibodies, can be first noted by day 1 and becomes associated with the lesions by 5 days postinoculation. These lesions become necrotic, with some evidence of in situ apoptosis. The lesions do not contain B, T, or NK cells. Rather, the lesions are largely composed of two subpopulations of Mac-1⁺ cells that are associated with the bacteria. Gr-1⁺ Mac-1⁺ immature myeloid cells and major histocompatibility complex class II-positive (MHC-II⁺) Mac-1⁺ macrophages were the most abundant cell phenotypes found in the granuloma and are likely major contributors in controlling the infection in its early stages. Our findings have shown that there is an early development of hepatic lesions where F. tularensis colocalizes with both Gr-1⁺ Mac-1⁺ and MHC-II⁺ Mac-1⁺ cells.

Top Abstract Introduction Materials and Methods Results and Discussion References

 
Francisella tularensis is a facultative intracellular bacterial zoonotic pathogen that causes a disease known as tularemia. Human infection can follow ingestion of contaminated food or water, contact of open skin wounds with infected animal carcasses, bites from various blood-sucking arthropods, or inhalation of aerosolized bacteria (2, 17, 18). In its natural setting, tularemia is the third most common tick-borne disease; it is also the second most common laboratory-associated infection in the United States (25, 39). Although tularemia has declined steadily since World War II, interest in Francisella continues not only as a model of study for intracellular bacteria but also due to its potential use as a biological weapon (16).

Two subspecies of F. tularensis, F. tularensis subsp. tularensis (type A) and F. tularensis subsp. holarctica (type B), are highly infectious in humans. Type B strains cause only moderate illness and are usually nonfatal. Meanwhile, type A strains cause potentially lethal infections in humans, particularly following exposure to aerosolized organisms. For this reason, type A F. tularensis is considered a potential biological warfare agent (16) and has been classified as a category A agent of bioterrorism by the Centers for Disease Control and Prevention. An attenuated live vaccine strain (LVS) derived from type B F. tularensis does not cause illness in humans but causes a disease in mice that resembles human tularemia (3, 21). Therefore, the LVS strain has been used extensively for experimental studies on the pathogenesis of tularemia. The involvement of the liver in both clinical and experimental tularemia regardless of the portal of entry or host species has been known for a long time (5, 18, 42, 43).

Single or multiple randomly distributed irregular microabscesses of mononuclear cells and a few neutrophils in the hepatic parenchyma have been seen as early as 1 day postinoculation (DPI) in murine tularemia (13). These microabscesses grow into well-circumscribed granulomas composed mostly of macrophages by 4 to 5 DPI. Hepatocytes can be infected by F. tularensis, and these cells can harbor large numbers of bacteria (11-13, 15, 33; H. Zheng and M. B. Furie, unpublished observations). With time, the developing granulomas become prominent in the entire liver, and the cytoplasm of many hepatocytes becomes completely filled with bacteria (15). Liver infection from LVS has also been used to study protective immunity and mouse strain susceptibility (12). Livers from LVS-immunized C57BL/6 mice contained small- to medium-sized areas of focal inflammatory necrosis with both necrotic and apoptotic hepatocytes, while the liver pathology of LVS-immunized BALB/c mice was milder. This mouse strain was more resistant to intradermal and aerosol inoculation (12). Thus, in murine tularemia, pathogen virulence, genetic background of the host, and route of inoculation all play a role in pathogenesis, specifically in the liver.

While the liver pathology of tularemia is well recognized in a number of experimental models, characterization of the infiltrating cells of the lesions has not been done with specific markers, nor, for that matter, has the process of cell death in liver infection been documented specifically. In this study, we used experimental sublethal tularemia infection of C3H/HeN mice to characterize the liver infiltrates and other signs of hepatic dysfunction. We report that subpopulations of cells expressing Mac-1 associate with F. tularensis during the early development of hepatic lesions.

Top Abstract Introduction Materials and Methods Results and Discussion References

 
Bacteria. F. tularensis LVS (ATCC 29684; American Type Culture Collection, Manassas, VA) was cultured in Mueller-Hinton (MH) broth (BD Biosciences, Sparks, MD) supplemented with 2% IsoVitaleX Enrichment (BD Biosciences), 0.1% glucose, 63 mM CaCl₂, 53 mM MgCl₂, and 34 mM ferric pyrophosphate and incubated at 37°C in 5% carbon dioxide. Mid-log-phase bacteria were frozen in 1-ml aliquots at –80°C (20, 24). Bacteria from frozen aliquots were grown on Chocolate II agar (BD Biosciences) at 37°C in 5% carbon dioxide. Single colonies were inoculated into prewarmed (37°C) MH broth, grown for 16 to 18 h, serially diluted in MH broth to 10⁵ CFU/ml as determined by an optical density reading at 600 nm, and verified by growth on Chocolate II agar.

Mice. Female C3H/HeN mice were purchased from Charles River Laboratories (Wilmington, MA) and used from 6 to 10 weeks of age. All mice were housed in microisolator cages with free access to food and water. Mice received intradermal injections of 10⁵ CFU of F. tularensis LVS. At various time points postinoculation, mice were euthanized, which was immediately followed by blood and organ collection. All animal procedures were approved by an institutional review board. The number of viable bacteria in blood was determined by streaking samples onto Chocolate II agar plates and counting the numbers of colonies.

White blood cell counts and enzymes. Total white blood cell counts were done manually by use of Petroff-Hausser chambers. Differentials were determined by enumeration from Giemsa-stained peripheral blood smears. Serum clinical chemistries for liver and kidney function were done by the Research Animal Diagnostic Laboratory, Columbia, MO. The tests included determinations for alanine transferase (ALT), alkaline phosphatase, direct and total bilirubin, lactate dehydrogenase (LDH), creatinine, and blood urea nitrogen.

Cell isolation. Following euthanization of mice, livers were perfused with large volumes of Hanks' balanced salt solution (Invitrogen, Grand Island, NY) until the organ was blanched. Once removed, livers were minced and incubated in digestive medium (0.05% collagenase A [Roche, Indianapolis, IN] and 0.002% DNase I [Sigma, St. Louis, MO] in Hanks' balanced salt solution) at 37°C and at 80 rpm for 30 min to provide a single-cell suspension of tissue. Cells were collected and centrifuged for 10 min at 400 x g followed by suspension on a Percoll gradient (GE Healthcare, Piscataway, NJ) and centrifugation for 30 min at room temperature (RT) at 400 x g in a swing-out rotor. Mononuclear cells were enumerated by using Petroff-Hausser chambers prior to antibody staining for flow cytometry.

Flow cytometry. Mononuclear cells (10⁶ cells) were resuspended in fluorescence-activated cell sorter buffer (0.2% bovine serum albumin [Sigma] and 0.09% NaN₃ [Sigma] in phosphate-buffered saline [PBS] [Invitrogen]) and incubated with anti-FcR antibody (clone 2.4G2) (BD Pharmingen, San Diego, CA) before appropriate amounts of conjugated antibodies or isotype controls were added and incubated for 30 min at 4°C (see below). Cells were washed twice with fluorescence-activated cell sorter buffer and centrifuged for 5 min at 400 x g at 4°C before being fixed in 500 µl 1% formalin in PBS. At least 10,000 viable cells were acquired on the basis of forward light and side light scattering and then quantified by using a BD FACSCalibur instrument and analyzed with WinList software (Verity Software House, Topsham, ME). Two-tailed P values were calculated using an unpaired t test with InStat software (GraphPad, San Diego, CA).

Antibodies for flow cytometry and immunofluorescence. The following antibodies were used for flow cytometry and confocal microscopy: fluorescein isothiocyanate (FITC) anti-mouse CD45R/B220 (clone RA3-6B2), FITC anti-mouse CD11c (clone HL3), FITC anti-mouse CD49b/Pan natural killer (NK) cells (clone DX5), R-phycoerythrin (PE) anti-mouse CD3 (clone 17A2), PE anti-mouse CD45R/B220 (clone RA3-6B2), PE anti-mouse CD11c (clone HL3), PE anti-mouse I-A/I-E (major histocompatibility complex class II [MHC-II]) (clone M5/114.15.2), PE anti-mouse Ly-6G and Ly-6C (Gr-1) (clone RB6-8C5), peridinin chlorophyll a protein (PerCP) anti-mouse CD4 (clone RM4-5), PerCP-Cy5.5 anti-mouse Mac-1 (CD11b) (clone M1/70), allophycocyanin (APC) anti-mouse NK1.1 (clone PK136), and APC anti-mouse CD8 (clone 53-6.7) from BD Pharmingen; Alexa Fluor 488 anti-mouse CD4 (clone GK1.5), Alexa Fluor 647 anti-mouse CD8a (clone 53-6.7), and Alexa Fluor 647 anti-mouse Mac-1 (CD11b) (clone M1/70) from Biolegend (San Diego, CA); and Alexa Fluor 488 anti-mouse F4/80 (clone CI:A3-1) from Serotec (Raleigh, NC). Isotype-matched antibodies (all from BD Pharmingen) were used as controls for nonspecific binding. Polyclonal rabbit anti-F. tularensis LVS was harvested after four injections of heat-killed organisms. FITC anti-rabbit immunoglobulin G (IgG) from Chemicon Int. (Temecula, CA) or Alexa Fluor 555 anti-rabbit IgG from Molecular Probes (Eugene, OR) was used as a secondary antibody to F. tularensis antisera.

Hematoxylin and eosin staining and immunohistology on tissue sections. Livers were aseptically removed and immediately fixed in 10% neutral buffered formalin, embedded in Blue Ribbon paraffin (Surgipath, Richmond, IL), sectioned at 5 µm, stained with hematoxylin and eosin, dehydrated in graded alcohols, cleared with xylene, and mounted with Acrymount (Statlab Medical Products, Lewisville, TX). Tissue sections were examined by light microscopy.

Detection of caspase-3 was achieved by dewaxing and rehydration of paraffin sections with xylene and graded alcohols, followed by quenching of endogenous peroxidase with methanol and hydrogen peroxide and blocking with Tween-bovine serum albumin. Rabbit anti-cleaved caspase-3 (Asp175) from Cell Signaling Technology (Danvers, MA) was diluted in blocking solution and added to sections for overnight incubation at RT. Sections were then washed and treated with polyclonal biotinylated anti-goat IgG (Vector Laboratories, Burlingame, CA) for 1 h at RT. Sections were washed, and avidin-biotinylated enzyme complex reagent (Vector Laboratories) was added for 45 min at RT, followed by five washes and incubation with diaminobenzidine (Sigma-Aldrich Corporation, St. Louis, MO) for 10 min. Sections were rinsed in water, counterstained with hematoxylin, dehydrated in graded alcohols, and cleared with xylene.

Terminal deoxynucleotidyl transferase-mediated dUTP-X nick end labeling (TUNEL) assays were performed according to the manufacturer's protocol using an in situ cell death detection kit with tetramethylrhodamine red (Roche Applied Science, Indianapolis, IN).

For detection of bacteria in the liver, paraffin sections were treated for 30 min with rabbit anti-F. tularensis LVS IgG after dewaxing and rehydration of the sections. Secondary alkaline phosphatase-labeled goat anti-rabbit IgG from Zymed (San Francisco, CA) was added for 30 min at RT, and Vulcan Fast red chromogen (Biocarta, San Diego, CA) was then used to visualize the bacteria.

Immunofluorescent staining of frozen tissue sections. Tissues removed from mice were immediately placed into freshly made 1% formalin in PBS from Invitrogen and gently shaken for 1 h at 4°C. The tissues were removed, blotted dry, placed into freshly made 30% sucrose in PBS at 4°C, and left overnight. The tissues were removed, blotted dry, placed into Neg –50 freezing compound (Richard-Allan Scientific, Kalamazoo, MI), rapidly frozen in isopentane that had been cooled with liquid nitrogen, and stored at –80°C. For some experiments, organs were immersed in OCT compound (Sakura Finetek, Torrance, CA) and then frozen and stored as described above.

Frozen tissue sections were cut at 5 µm in the cryostat at –25°C, air dried, and fixed in acetone for 30 s. Twenty microliters of the various antibodies (see above) diluted in 0.01 M PBS (pH 7.4) was applied to sections and incubated in the dark for 25 min. Slides were washed three times in PBS, and when appropriate, secondary antibodies were added for 25 min in the dark. Mouse spleens, treated in the same manner, were used as positive controls for the antibodies used in this study. After washing, slides were mounted in Opti-Mount (Richard-Allan Scientific, Kalamazoo, MI). The slides were examined by phase-contrast and epifluorescence microscopy using a Nikon Eclipse E600 microscope, and images were captured using a Spot camera (Diagnostic Instruments, Inc.). Slides for confocal microscopy were analyzed using a Leica DM IRE2 confocal microscope. Images of the red, green, and blue emission signals were captured separately with the Leica LCS software package. Images were processed using Adobe Photoshop.

Top Abstract Introduction Materials and Methods Results and Discussion References

 
The pathology of liver involvement in experimental tularemia has been studied with standard histopathological procedures, and the existence of granulomatous necrotic lesions has been noted previously in several studies (13, 15). To date, however, the infiltrating cells of the hepatic lesions have not been characterized with respect to specific markers, and the extent of apoptosis has not been examined with markers specific to this type of cell death.

Intradermal inoculation of C3H/HeN mice with F. tularensis LVS led to bacteremia for the first 5 DPI. In the periphery, this bacteremia was accompanied by leukocytosis with an initial reversal in the ratio of lymphocytes to neutrophils in the differential as well as a modest increase in the percentage of circulating monocytes at 4 DPI (Table 1). A similar pattern of leukocytosis and reversal of the differential has been demonstrated for experimental infections of other strains of mice (13). There were marked increases in serum levels of ALT (Fig. 1A) and LDH (Fig. 1B). This pattern is consistent with early inflammation in the liver without reducing the ability of the liver to conjugate and secrete bilirubin, as evidenced by the normal values obtained for direct and indirect bilirubin and alkaline phosphatase (data not shown). Kidney function was within normal limits.

View this table: [in this window] [in a new window]

TABLE 1. Total and differential peripheral blood leukocyte counts from mice inoculated with F. tularensis LVS

View larger version (10K): [in this window] [in a new window]

FIG. 1. Mean serum levels ± standard deviations of (A) ALT and (B) LDH in mice infected with sublethal doses of LVS. The dashed line represents the upper limit of the normal range for each enzyme. Each diamond represents the mean value ± standard deviation for three mice.

Here, we confirm the previous findings of widespread, early foci characterized by the infiltration of a large number of mononuclear cells that are morphologically consistent with macrophages and a few neutrophils in the liver (Fig. 2B) (12, 13). These lesions had a focus of mononuclear infiltration. As these lesions matured, necrotic hepatocytes with pyknotic nuclei were common within the inflammatory foci (Fig. 2C and D). The evolution of the granulomatous response is typical, where neutrophils with a short half-life appear early and mononuclear cells persist. Neutrophils are known to be important for defense in primary tularemia infection (14, 40). The perivascular location of many of the granulomas (Fig. 2C) suggests that the infiltrate derives from circulating cells from the blood as opposed to an expansion of the resident cells.

View larger version (144K): [in this window] [in a new window]

FIG. 2. Hematoxylin- and eosin-stained sections of livers of mice infected with sublethal doses of LVS. (A) Normal liver. (B) Low-power view of hepatic parenchyma showing multiple lesions (arrows) of inflammatory cell infiltrates in the liver of a mouse inoculated with LVS 5 days earlier. (C) Mononuclear cell inflammatory cell infiltrate of a perivascular focus of inflammation in the liver of a mouse infected with LVS 5 days earlier. Some neutrophils are evident, as indicated by arrows. (D) A 5-day-old lesion within the hepatic parenchyma showing necrotic hepatocytes, pyknotic nuclei, and residual mononuclear cell infiltrate. Bars, 150 µm (A and B) and 75 µm (C and D).

Bacteria invade the liver parenchyma early in randomly distributed locations (Fig. 3B). Some bacteria appear to be associated with Kupffer cells based on the location of these cells on the sinusoids, but others are within hepatocytes (Fig. 3B and C). Figure 3C shows a hepatocyte swollen with bacteria, similar to what has been observed previously by others (15). These heavily infected hepatocytes could become focal points for the development of the granulomas. In the granuloma at 5 DPI, it is difficult to determine whether the bacteria are extracellular or associated with hepatocytes, macrophages, or both (Fig. 3D). Nonetheless, there is severe damage to hepatocytes, and numerous bacterial colonies are present in the lesions. Our results indicated that hepatic dysfunction in tularemia is likely to be a contributor to the morbidity and mortality of this infection, although in some instances, liver disease can be reversible.

View larger version (164K): [in this window] [in a new window]

FIG. 3. Detection of bacteria in livers of mice inoculated with sublethal doses of LVS 5 days earlier. Bacteria (red) were detected with rabbit anti-F. tularensis LVS followed by alkaline phosphatase-conjugated anti-rabbit serum. Vulcan Fast red was used to detect alkaline phosphatase. (A) Normal liver. (B) Low-power view of hepatic parenchyma showing multiple sites of bacterial infection. (C) Hepatocytes infected with F. tularensis. Some of the hepatocytes are heavily infected and swollen with bacteria. (D) Bacteria concentrated within the granuloma. Bars, 150 µm (A and B) and 50 µm (C and D).

To characterize the cells in the lesions observed in the livers of infected mice, immunofluorescence microscopy of frozen liver sections was performed using specific cell surface markers for macrophages, lymphocytes, and F. tularensis antibodies. Infection of the liver parenchyma was already present on the first day after inoculation (Fig. 4). In later stages, the vast majority of the mononuclear cells within the granulomas were F4/80^– Mac-1⁺ (Fig. 5). A few F4/80⁺ Mac-1^– cells were found in the borders of the lesion; these cells may represent a population of Kupffer cells (28). The F4/80^– Mac-1⁺ cells may represent monocytes/macrophages recruited from the blood to the liver. Cells with this phenotype have been shown to traffic from the peripheral blood to the inflamed retina in a murine model of autoimmune uveoretinitis (45). Blood monocytes can express both F4/80 and Mac-1 markers (23), so it is possible that the phenotype of the infiltrating mononuclear cells may be derived from blood, with a subsequent downregulation of the F4/80 marker. Another possibility is that F4/80^– Mac-1⁺ cells are a subset of resident macrophages similar to those in the spleen, optic nerve, and the connective tissue of the lung in which the expression of the F4/80 antigen is downregulated by inflammatory stimuli (6, 8, 19, 44). Regardless of the possible origin of Mac-1⁺ F4/80^– cells, F. tularensis was found to be associated with these cells, correlating with a Mac-1⁺ phenotype that has been shown to be involved in the hepatic killing of other bacteria (7).

View larger version (44K): [in this window] [in a new window]

FIG. 4. Immunofluorescent detection of F. tularensis and inflammatory cells in the livers of mice inoculated with sublethal doses 1 day earlier viewed by confocal microscopy. (A) Merged-image, low-power view of Mac-1+ cells (Alexa Fluor 647, blue), F. tularensis (secondary anti-rabbit Ig, Alexa Fluor 555, red), and F4/80 (Alexa Fluor 488, green). (B) Hepatocytes infected with F. tularensis (red). (C) Merged image of a Mac-1+ cell (neutrophil-like, blue) infected with F. tularensis (red).

View larger version (14K): [in this window] [in a new window]

FIG. 5. Immunofluorescent detection of F. tularensis and inflammatory cells in the livers of mice inoculated with sublethal doses 5 days earlier viewed by confocal microscopy. (A) F4/80+ cells (Alexa Fluor 488, green). (B) F. tularensis (secondary anti-rabbit Ig, Alexa Fluor 555, red) in foci of inflammatory cells. (C) Mac-1+ cells (Alexa Fluor 647, blue). (D) Merged image of panels A to C. Note the colocalization of Mac-1+ cells (blue) and F. tularensis (red). Also note the peripheral location of the F4/80+ cells around the foci of infection. Some cells are both F4/80+ and Mac-1+ and can be seen in merged images of panels E and F.

Mac-1 can be expressed on a variety of cells, including granulocytes, T cells, B cells, NK cells, dendritic cells (DCs), and monocytes. To further characterize the Mac-1⁺ infiltrating mononuclear cells, immunofluorescence microscopy was performed using specific markers for cell types known to express Mac-1. Neutrophils were ruled out as a major cell type contributing to the infiltrate of the granulomas by morphology (Fig. 2C and D). CD3⁺, CD4⁺, and CD8⁺ T cells and B220⁺ B cells were not detected in the granulomatous areas of hepatocyte necrosis at 1 and 5 DPI (data not shown). However, the Mac-1⁺ cells did colocalize with markers specific for myeloid cell populations that were most consistent with macrophages and DCs. One population expressed both macrophage (Mac-1⁺) and granulocyte (Gr-1⁺) markers and was the predominant phenotype in the granuloma (Fig. 6A). The lesions also contained a significant population that had an MHC-II⁺ Mac-1⁺ phenotype (Fig. 6B) and a CD11c⁺ Mac-1⁺ DC phenotype (Fig. 6C). NK cells (NK1.1⁺ and CD49/DX5⁺) were seen in the liver tissue at 5 DPI but were not associated with the hepatic lesions (Fig. 6D).

View larger version (22K): [in this window] [in a new window]

FIG. 6. Immunofluorescent detection of F. tularensis and inflammatory cells in the livers of mice inoculated with sublethal doses 5 days earlier viewed by confocal microscopy. (A) Merged image of Mac-1+ cells (Alexa Fluor 647, blue), Gr-1+ cells (PE, red), and F. tularensis (secondary anti-rabbit Ig, FITC, green). Gr-1+ cells colocalize with Mac-1+ cells (pink) and make up the granuloma. (B) Merged image of Mac-1+ cells (Alexa Fluor 647, blue), MHC-II+ cells (PE, red) and F. tularensis (secondary anti-rabbit Ig, FITC, green). The granuloma also consists of a cell population, MHC II+ Mac-1+, that also associates with the bacteria. (C) Merged image of Mac-1+ cells (Alexa Fluor 647, blue), CD11c+ cells (PE, red), and F. tularensis (secondary anti-rabbit Ig, FITC, green). DCs (CD11c+ Mac-1+) were found in association with the granuloma. (D) Merged image of NK1.1+ cells (APC, blue), F. tularensis (secondary anti-rabbit Ig, Alexa Fluor 555, red), and DX5+ (FITC, green). NK cells were seen in the tissue but not in association with the bacteria or the hepatic lesions.

Coexpression of both Gr-1 and Mac-1 is indicative of an immature myeloid cell that can differentiate into either a mature granulocyte, macrophage, or DC (for a review, see reference 38). Immature myeloid cells (Gr-1⁺ Mac-1⁺) do not express MHC-II molecules and exhibit dull F4/80 expression (29, 34), correlating with the predominant phenotype (F4/80^– Mac-1⁺) seen at 5 DPI (Fig. 5). Also known as myeloid suppressor cells, these cells accumulate and inhibit the T-cell immune response in tumor-bearing mice (22, 36, 41). In addition, they have been found to have immunosuppressive effects in mice infected with various pathogens (1, 26, 35). However, it has been noted that depending on the cytokine milieu that is present, Gr-1⁺ Mac-1⁺ cells can either activate or inactivate the T-lymphocyte immune response. Bronte et al. (9) previously showed that when cultured in vitro with proinflammatory cytokines (gamma interferon and tumor necrosis factor alpha), Gr-1⁺ Mac-1⁺ cells differentiated into functional antigen-presenting cells. However, when these cells were cultured with an anti-inflammatory cytokine (interleukin-4), the cells greatly increased T-cell suppression. Therefore, the function of immature myeloid cells is dependent upon the host inflammatory response initiated by a pathological process.

To further quantify the abundance of cellular populations, flow cytometry analysis was performed on liver tissue from uninfected mice and mice infected with F. tularensis LVS at 5 DPI. Markers were used for T cells (CD3), B cells (B220), DCs (CD11c and Mac-1), NK cells (DX5 and NK1.1), macrophages (F4/80, MHC-II, and Mac-1), and immature myeloid cells (Gr-1 and Mac-1) (Table 2). Results of the quantification of mononuclear cells in the liver by flow cytometry were consistent with the imaging results. Markers for B and T cells were similarly expressed in both uninfected mice and mice inoculated with F. tularensis LVS 5 days earlier. A twofold increase of NK cell marker expression by 5 DPI was noted. This may indicate that NK cells are upregulated to aid the innate response and cytokine secretion, even though NK cells were not seen in the granulomas. Total CD11c expression, which is indicative of DCs, increased 2.5-fold by 5 DPI. Furthermore, the numbers of myeloid DCs (CD11c⁺ Mac-1⁺), which were found in the lesions, also increased but did not reach statistical significance. The most significant increase of all cellular phenotypes was Mac-1⁺ cells (2.3% uninfected to 21.3% at 5 DPI), which correlates to the majority of cells seen in the hepatic lesions. The bulk of Mac-1⁺ cells were CD11c^–, indicating that most Mac-1⁺ cells were not DCs. In addition, levels of F4/80⁺ Mac-1⁺ cells did not increase significantly, correlating with the low levels of this subpopulation shown in Fig. 5.

View this table: [in this window] [in a new window]

TABLE 2. Flow cytometry analysis of cell marker expression of total cell counts from livers of mice inoculated with F. tularensis LVS 5 days earlier compared to that from uninfected livers

The largest increases in Mac-1⁺ subpopulations observed by flow cytometry analysis were the Gr-1⁺ Mac-1⁺ immature myeloid cells and the MHC-II⁺ Mac-1⁺ CD11c^– macrophages, confirming the results from immunofluorescence staining of liver tissue. Representative plots from flow cytometry are shown in Fig. 7A and B. Correlating to their abundance in the granulomas, Gr-1⁺ Mac-1⁺ and MHC-II⁺ Mac-1⁺ CD11c^– cells are likely major contributors in controlling early F. tularensis LVS infection. The Gr-1⁺ Mac-1⁺ cells could function as immunosuppressive cells to inhibit the immune response and allow for bacterial survival or as a means to wall off the infection until an inflammatory response develops. It is tempting to speculate that under a proinflammatory response, Gr-1⁺ Mac-1⁺ cells differentiate into functional antigen-presenting cells largely as MHC-II⁺ Mac-1⁺ CD11c^– macrophages and to a lesser extent as myeloid DCs (CD11c⁺ Mac-1⁺). These cell phenotypes would correlate with those seen within the hepatic lesions (Fig. 6).

View larger version (26K): [in this window] [in a new window]

FIG. 7. Flow cytometry analysis of predominant cell populations in the livers of F. tularensis LVS-infected mice. Accumulation of Gr-1+ Mac-1+ immature myeloid cells (A) and MHC-II+ Mac-1+ cell expression (B) in a mouse 5 DPI compared to uninfected control are shown. A representative experiment is shown for both A and B.

We have observed and confirmed a necrotic process (12, 13) that is clearly evident within the liver abscesses at 5 DPI (Fig. 2). Apoptosis of hepatocytes is the hallmark of murine listeriosis, which is caused by another intracellular organism with a predilection for liver involvement (37). In addition, a number of in vitro studies with murine macrophages have shown that F. tularensis infection is able to trigger the apoptotic cascade (27, 30-32). Based on those studies, we examined the extent of apoptosis in the livers of infected mice. Although apoptotic cells were detected in the livers of mice infected with F. tularensis using both caspase-3 and TUNEL markers (Fig. 8), these levels of apoptosis were qualitatively less than those observed in a sublethal Listeria infection (37, 46). Both the necrotic and apoptotic pathways of cell death appear to be important in tularemia, as is also true for some other bacterial infections of the liver. For example, Listeria induces the apoptosis of hepatocytes (37), but it is also known that murine macrophages succumb to Listeria infection in vitro by necrosis (4). Furthermore, apoptotic death of CD8⁺ T cells has been attributed to a function of Gr-1⁺ Mac-1⁺ cells (10). Infection of the liver by microorganisms results in the death of hepatocytes, and myeloid cells could be major contributors to this process.

View larger version (91K): [in this window] [in a new window]

FIG. 8. Detection of apoptotic cell markers by immunohistochemistry in the livers of mice inoculated with sublethal doses of F. tularensis 5 days earlier. (A) Low-power view of anti-cleaved caspase-3 reactivity (brown) within the foci of inflammation and infection. (B and C) High-power view of anti-cleaved caspase-3 reactivity (brown) within the foci of inflammation and infection. (D) Low-power phase-contrast view of liver parenchyma containing foci of inflammation and infection. (E) TUNEL-positive cells (red) (same field as panel D). (F) Merged photograph of panels D and E showing cells with fragmented DNA. Bars, 150 µm (A) and 75 µm (B to G).

F. tularensis causes liver damage, as evidenced by elevated serum ALT and LDH levels and by the granulomatous-necrotic-apoptotic lesions that appear by 5 DPI. These lesions are composed mostly of Mac-1⁺ cells from two myeloid populations (Gr-1⁺ Mac-1⁺ and MHC-II⁺ Mac-1⁺) that are associated with the bacteria, thus suggesting that these cells are important for controlling the infection. These cells appear to be recruited cells and are accumulated specifically to regulate the infection in its early stage and may therefore prove useful in the development of vaccines against F. tularensis infections to stimulate the activation of these particular cell subsets.

 
This work was supported by a grant from the National Institutes of Health, AI 055621.

We appreciate the assistance of Gloria Monsalve and Patricio Mena.

 
* Corresponding author. Mailing address: Center for Infectious Diseases, 5120 Centers for Molecular Medicine, Stony Brook, NY 11794-5120. Phone: (631) 632-4225. Fax: (631) 632-4294. E-mail: jbenach{at}notes.cc.sunysb.edu.

Published ahead of print on 25 September 2006.

Editor: J. T. Barbieri

Present address: Laboratorio de Espiroquetas y Patógenos Especiales, Centro Nacional de Microbiología, Instituto de Salud Carlos III, Majadahonda 28220, Spain.

Top Abstract Introduction Materials and Methods Results and Discussion References

 

1
1. al-Ramadi, B. K., M. A. Brodkin, D. M. Mosser, and T. K. Eisenstein. 1991. Immunosuppression induced by attenuated Salmonella. Evidence for mediation by macrophage precursors. J. Immunol. 146:2737-2746.[Abstract]
2
2. Anda, P., J. Segura del Pozo, J. M. Diaz García, R. Escudero, F. J. Garcí Peña, M. C. López Velasco, R. E. Sellek, M. R. Jiménez Chillarón, L. P. Sánchez Serrano, and J. F. Martínez Navarro. 2001. Waterborne outbreak of tularemia associated with crayfish fishing. Emerg. Infect. Dis. 7:575-582.[Medline]
3
3. Anthony, L. S., and P. A. Kongshavn. 1987. Experimental murine tularemia caused by Francisella tularensis, live vaccine strain: a model of acquired cellular resistance. Microb. Pathog. 2:3-14.[CrossRef][Medline]
4
4. Barsig, J., and S. H. Kaufmann. 1997. The mechanism of cell death in Listeria monocytogenes-infected murine macrophages is distinct from apoptosis. Infect. Immun. 65:4075-4081.[Abstract]
5
5. Baskerville, A., and P. Hambleton. 1976. Pathogenesis and pathology of respiratory tularaemia in the rabbit. Br. J. Exp. Pathol. 57:339-347.[Medline]
6
6. Breel, M., M. Van der Ende, T. Sminia, and G. Kraal. 1988. Subpopulations of lymphoid and non-lymphoid cells in bronchus-associated lymphoid tissue (BALT) of the mouse. Immunology 63:657-662.[Medline]
7
7. Brengman, M. L., D. Wang, K. B. Wilkins, N. Sakamoto, T. Arai, E. P. Ceppa, A. S. Klein, and G. B. Bulkley. 2003. Hepatic killing but not clearance of systemically circulating bacteria is dependent upon peripheral leukocytes via Mac-1 (CD11b/CD18). Shock 19:263-267.[CrossRef][Medline]
8
8. Brissette-Storkus, C. S., S. M. Reynolds, A. J. Lepisto, and R. L. Hendricks. 2002. Identification of a novel macrophage population in the normal mouse corneal stroma. Investig. Ophthalmol. Vis. Sci. 43:2264-2271.[Abstract/Free Full Text]
9
9. Bronte, V., E. Apolloni, A. Cabrelle, R. Ronca, P. Serafini, P. Zamboni, N. P. Restifo, and P. Zanovello. 2000. Identification of a CD11b(+)/Gr-1(+)/CD31(+) myeloid progenitor capable of activating or suppressing CD8(+) T cells. Blood 96:3838-3846.[Abstract/Free Full Text]
10
10. Bronte, V., M. Wang, W. W. Overwijk, D. R. Surman, F. Pericle, S. A. Rosenberg, and N. P. Restifo. 1998. Apoptotic death of CD8+ T lymphocytes after immunization: induction of a suppressive population of Mac-1+/Gr-1+ cells. J. Immunol. 161:5313-5320.[Abstract/Free Full Text]
11
11. Chen, W., R. KuoLee, H. Shen, and J. W. Conlan. 2004. Susceptibility of immunodeficient mice to aerosol and systemic infection with virulent strains of Francisella tularensis. Microb. Pathog. 36:311-318.[CrossRef][Medline]
12
12. Chen, W., H. Shen, A. Webb, R. KuoLee, and J. W. Conlan. 2003. Tularemia in BALB/c and C57BL/6 mice vaccinated with Francisella tularensis LVS and challenged intradermally, or by aerosol with virulent isolates of the pathogen: protection varies depending on pathogen virulence, route of exposure, and host genetic background. Vaccine 21:3690-3700.[CrossRef][Medline]
13
13. Conlan, J. W., W. Chen, H. Shen, A. Webb, and R. KuoLee. 2003. Experimental tularemia in mice challenged by aerosol or intradermally with virulent strains of Francisella tularensis: bacteriologic and histopathologic studies. Microb. Pathog. 34:239-248.[CrossRef][Medline]
14
14. Conlan, J. W., R. KuoLee, H. Shen, and A. Webb. 2002. Different host defences are required to protect mice from primary systemic vs pulmonary infection with the facultative intracellular bacterial pathogen, Francisella tularensis LVS. Microb. Pathog. 32:127-134.[CrossRef][Medline]
15
15. Conlan, J. W., and R. J. North. 1992. Early pathogenesis of infection in the liver with the facultative intracellular bacteria Listeria monocytogenes, Francisella tularensis, and Salmonella typhimurium involves lysis of infected hepatocytes by leukocytes. Infect. Immun. 60:5164-5171.[Abstract/Free Full Text]
16
16. Dennis, D. T., T. V. Inglesby, D. A. Henderson, J. G. Bartlett, M. S. Ascher, E. Eitzen, A. D. Fine, A. M. Friedlander, J. Hauer, M. Layton, S. R. Lillibridge, J. E. McDade, M. T. Osterholm, T. O'Toole, G. Parker, T. M. Perl, P. K. Russell, and K. Tonat. 2001. Tularemia as a biological weapon: medical and public health management. JAMA 285:2763-2773.[Abstract/Free Full Text]
17
17. Ellis, J., P. C. Oyston, M. Green, and R. W. Titball. 2002. Tularemia. Clin. Microbiol. Rev. 15:631-646.[Abstract/Free Full Text]
18
18. Evans, M. E., D. W. Gregory, W. Schaffner, and Z. A. McGee. 1985. Tularemia: a 30-year experience with 88 cases. Medicine (Baltimore) 64:251-269.[Medline]
19
19. Ezekowitz, R. A., J. Austyn, P. D. Stahl, and S. Gordon. 1981. Surface properties of bacillus Calmette-Guerin-activated mouse macrophages. Reduced expression of mannose-specific endocytosis, Fc receptors, and antigen F4/80 accompanies induction of Ia. J. Exp. Med. 154:60-76.[Abstract/Free Full Text]
20
20. Forestal, C. A., J. L. Benach, C. Carbonara, J. K. Italo, T. J. Lisinski, and M. B. Furie. 2003. Francisella tularensis selectively induces proinflammatory changes in endothelial cells. J. Immunol. 171:2563-2570.[Abstract/Free Full Text]
21
21. Fortier, A. H., M. V. Slayter, R. Ziemba, M. S. Meltzer, and C. A. Nacy. 1991. Live vaccine strain of Francisella tularensis: infection and immunity in mice. Infect. Immun. 59:2922-2928.[Abstract/Free Full Text]
22
22. Gabrilovich, D. I., M. P. Velders, E. M. Sotomayor, and W. M. Kast. 2001. Mechanism of immune dysfunction in cancer mediated by immature Gr-1+ myeloid cells. J. Immunol. 166:5398-5406.[Abstract/Free Full Text]
23
23. Geissmann, F., S. Jung, and D. R. Littman. 2003. Blood monocytes consist of two principal subsets with distinct migratory properties. Immunity 19:71-82.[CrossRef][Medline]
24
24. Gil, H., J. L. Benach, and D. G. Thanassi. 2004. Presence of pili on the surface of Francisella tularensis. Infect. Immun. 72:3042-3047.[Abstract/Free Full Text]
25
25. Goethert, H. K., I. Shani, and S. R. Telford III. 2004. Genotypic diversity of Francisella tularensis infecting Dermacentor variabilis ticks on Martha's Vineyard, Massachusetts. J. Clin. Microbiol. 42:4968-4973.[Abstract/Free Full Text]
26
26. Goñi, O., P. Alcaide, and M. Fresno. 2002. Immunosuppression during acute Trypanosoma cruzi infection: involvement of Ly6G (Gr1⁺)CD11b⁺ immature myeloid suppressor cells. Int. Immunol. 14:1125-1134.[Abstract/Free Full Text]
27
27. Hrstka, R., J. Stulík, and B. Vojtek. 2005. The role of MAPK signal pathways during Francisella tularensis LVS infection-induced apoptosis in murine macrophages. Microbes Infect. 7:619-625.[Medline]
28
28. Hume, D. A., I. L. Ross, S. R. Himes, R. T. Sasmono, C. A. Wells, and T. Ravasi. 2002. The mononuclear phagocyte system revisited. J. Leukoc. Biol. 72:621-627.[Abstract/Free Full Text]
29
29. Inaba, K., M. Inaba, M. Deguchi, K. Hagi, R. Yasumizu, S. Ikehara, S. Muramatsu, and R. M. Steinman. 1993. Granulocytes, macrophages, and dendritic cells arise from a common major histocompatibility complex class II-negative progenitor in mouse bone marrow. Proc. Natl. Acad. Sci. USA 90:3038-3042.[Abstract/Free Full Text]
30
30. Lai, X. H., I. Golovliov, and A. Sjöstedt. 2004. Expression of IglC is necessary for intracellular growth and induction of apoptosis in murine macrophages by Francisella tularensis. Microb. Pathog. 37:225-230.[Medline]
31
31. Lai, X. H., I. Golovliov, and A. Sjöstedt. 2001. Francisella tularensis induces cytopathogenicity and apoptosis in murine macrophages via a mechanism that requires intracellular bacterial multiplication. Infect. Immun. 69:4691-4694.[Abstract/Free Full Text]
32
32. Lai, X. H., and A. Sjöstedt. 2003. Delineation of the molecular mechanisms of Francisella tularensis-induced apoptosis in murine macrophages. Infect. Immun. 71:4642-4646.[Abstract/Free Full Text]
33
33. Lamps, L. W., J. M. Havens, A. Sjöstedt, D. L. Page, and M. A. Scott. 2004. Histologic and molecular diagnosis of tularemia: a potential bioterrorism agent endemic to North America. Mod. Pathol. 17:489-495.[CrossRef][Medline]
34
34. Marshall, M. A., D. Jankovic, V. E. Maher, A. Sher, and J. A. Berzofsky. 2001. Mice infected with Schistosoma mansoni develop a novel non-T-lymphocyte suppressor population which inhibits virus-specific CTL induction via a soluble factor. Microbes Infect. 3:1051-1061.[CrossRef][Medline]
35
35. Mencacci, A., C. Montagnoli, A. Bacci, E. Cenci, L. Pitzurra, A. Spreca, M. Kopf, A. H. Sharpe, and L. Romani. 2002. CD80+Gr-1+ myeloid cells inhibit development of antifungal Th1 immunity in mice with candidiasis. J. Immunol. 169:3180-3190.[Abstract/Free Full Text]
36
36. Otsuji, M., Y. Kimura, T. Aoe, Y. Okamoto, and T. Saito. 1996. Oxidative stress by tumor-derived macrophages suppresses the expression of CD3 zeta chain of T-cell receptor complex and antigen-specific T-cell responses. Proc. Natl. Acad. Sci. USA 93:13119-13124.[Abstract/Free Full Text]
37
37. Rogers, H. W., M. P. Callery, B. Deck, and E. R. Unanue. 1996. Listeria monocytogenes induces apoptosis of infected hepatocytes. J. Immunol. 156:679-684.[Abstract]
38
38. Serafini, P., C. De Santo, I. Marigo, S. Cingarlini, L. Dolcetti, G. Gallina, P. Zanovello, and V. Bronte. 2004. Derangement of immune responses by myeloid suppressor cells. Cancer Immunol. Immunother. 53:64-72.[CrossRef][Medline]
39
39. Shapiro, D. S., and D. R. Schwartz. 2002. Exposure of laboratory workers to Francisella tularensis despite a bioterrorism procedure. J. Clin. Microbiol. 40:2278-2281.[Abstract/Free Full Text]
40
40. Sjöstedt, A., J. W. Conlan, and R. J. North. 1994. Neutrophils are critical for host defense against primary infection with the facultative intracellular bacterium Francisella tularensis in mice and participate in defense against reinfection. Infect. Immun. 62:2779-2783.[Abstract/Free Full Text]
41
41. Song, X., Y. Krelin, T. Dvorkin, O. Bjorkdahl, S. Segal, C. A. Dinarello, E. Voronov, and R. N. Apte. 2005. CD11b+/Gr-1+ immature myeloid cells mediate suppression of T cells in mice bearing tumors of IL-1beta-secreting cells. J. Immunol. 175:8200-8208.[Abstract/Free Full Text]
42
42. Sutinen, S., and H. Syrjala. 1986. Histopathology of human lymph node tularemia caused by Francisella tularensis var palaearctica. Arch. Pathol. Lab. Med. 110:42-46.[Medline]
43
43. White, J. D., J. R. Rooney, P. A. Prickett, E. B. Derrenbacher, C. W. Beard, and W. R. Griffith. 1964. Pathogenesis of experimental respiratory tularemia in monkeys. J. Infect. Dis. 114:277-283.[Medline]
44
44. Witmer, M. D., and R. M. Steinman. 1984. The anatomy of peripheral lymphoid organs with emphasis on accessory cells: light-microscopic immunocytochemical studies of mouse spleen, lymph node, and Peyer's patch. Am. J. Anat. 170:465-481.[CrossRef][Medline]
45
45. Xu, H., A. Manivannan, R. Dawson, I. J. Crane, M. Mack, P. Sharp, and J. Liversidge. 2005. Differentiation to the CCR2+ inflammatory phenotype in vivo is a constitutive, time-limited property of blood monocytes and is independent of local inflammatory mediators. J. Immunol. 175:6915-6923.[Abstract/Free Full Text]
46
46. Zheng, S. J., P. Wang, G. Tsabary, and Y. H. Chen. 2004. Critical roles of TRAIL in hepatic cell death and hepatic inflammation. J. Clin. Investig. 113:58-64.[CrossRef][Medline]

---

Infection and Immunity, December 2006, p. 6590-6598, Vol. 74, No. 12
0019-9567/06/$08.00+0     doi:10.1128/IAI.00868-06
Copyright © 2006, American Society for Microbiology. All Rights Reserved.

This article has been cited by other articles:

• Wickstrum, J. R., Bokhari, S. M., Fischer, J. L., Pinson, D. M., Yeh, H.-W., Horvat, R. T., Parmely, M. J. (2009). Francisella tularensis Induces Extensive Caspase-3 Activation and Apoptotic Cell Death in the Tissues of Infected Mice. Infect. Immun. 77: 4827-4836 [Abstract] [Full Text]  
• Swirski, F. K., Weissleder, R., Pittet, M. J. (2009). Heterogeneous In Vivo Behavior of Monocyte Subsets in Atherosclerosis. Arterioscler. Thromb. Vasc. Bio. 29: 1424-1432 [Abstract] [Full Text]  
• Sharma, J., Li, Q., Mishra, B. B., Pena, C., Teale, J. M. (2009). Lethal pulmonary infection with Francisella novicida is associated with severe sepsis. J. Leukoc. Biol. 86: 491-504 [Abstract] [Full Text]  
• Collazo, C. M., Meierovics, A. I., De Pascalis, R., Wu, T. H., Lyons, C. R., Elkins, K. L. (2009). T Cells from Lungs and Livers of Francisella tularensis-Immune Mice Control the Growth of Intracellular Bacteria. Infect. Immun. 77: 2010-2021 [Abstract] [Full Text]  
• Savitt, A. G., Mena-Taboada, P., Monsalve, G., Benach, J. L. (2009). Francisella tularensis Infection-Derived Monoclonal Antibodies Provide Detection, Protection, and Therapy. CVI 16: 414-422 [Abstract] [Full Text]  
• Woolard, M. D., Hensley, L. L., Kawula, T. H., Frelinger, J. A. (2008). Respiratory Francisella tularensis Live Vaccine Strain Infection Induces Th17 Cells and Prostaglandin E2, Which Inhibits Generation of Gamma Interferon-Positive T Cells. Infect. Immun. 76: 2651-2659 [Abstract] [Full Text]  
• Bokhari, S. M., Kim, K.-J., Pinson, D. M., Slusser, J., Yeh, H.-W., Parmely, M. J. (2008). NK Cells and Gamma Interferon Coordinate the Formation and Function of Hepatic Granulomas in Mice Infected with the Francisella tularensis Live Vaccine Strain. Infect. Immun. 76: 1379-1389 [Abstract] [Full Text]  
• Lindemann, S. R., McLendon, M. K., Apicella, M. A., Jones, B. D. (2007). An In Vitro Model System Used To Study Adherence and Invasion of Francisella tularensis Live Vaccine Strain in Nonphagocytic Cells. Infect. Immun. 75: 3178-3182 [Abstract] [Full Text]  

This Article Abstract Full Text (PDF) Alert me when this article is cited Alert me if a correction is posted Services Similar articles in this journal Similar articles in PubMed Alert me to new issues of the journal Download to citation manager Reprints and Permissions Copyright Information Books from ASM Press MicrobeWorld Citing Articles Citing Articles via HighWire Citing Articles via Google Scholar Google Scholar Articles by Rasmussen, J. W. Articles by Benach, J. L. Search for Related Content PubMed PubMed Citation Articles by Rasmussen, J. W. Articles by Benach, J. L.